Wolbachia PCR
Table of Contents
Fall 2012
Wolbachia : Discover the M icrobes W ithin
Introduction…………………………………………………………………………………………………………..1
Laboratory Exercise: Insect Identification………….………………………………………………………….….5
Insect Identification Lab Report……………………………………………………………………………………7
Laboratory Exercise: DNA Extraction……….………………………………………………………………...….8
Important Laboratory Practices…………………………………………..…………………………..…..8
DNA Isolation from Insect Tissue……………………………………………………………………..….9
Laboratory Exercise: Polymerase Chain Reaction……………………………………………………...……..15
Illustration of the Polymerase Chain Reaction…………………………………………………………16
Setting up the Polymerase Chain Reaction…………………………………………………...……….17
Laboratory Exercise: Agarose Gel Electrophoresis………………………………………………………..…..19
Electrophoresis of Amplified DNA……………………………………………..………………………..20
Staining and Photographing Agarose Gels………………………………………………...…………..21
Instructions for using Invitrogen E-Gels……………………………..………………………………….22
Results…………………………………………………………………….………………………………………..24
Uploading Class Results to the BABEC Wolbachia Map…………………………………………………...…25
How to submit PCR samples for sequencing to CSU East Bay………………………………………………26
Bioinformatics Exercise: Sequence Taxonomy………………………………………………………………..28
Bioinformatics Exercise: Sequence Searching Using BLAST…………………….…………………………..31
Acknowledgements…………………………………………………………………..……………………………35
Wolbachia: Discover the Microbes Within
Nicole Markelz
BABEC
Katy Korsmeyer, Ph.D.
BABEC
The idea and methods for this lab are based on the materials developed by Dr. Seth Bordenstein and other
scientists of the Wolbachia Project at Woods Hole Marine Biological Laboratory (MBL). We gratefully acknowledge
Dr. Bordenstein, Michael Clark, Michele Bahr, and others at MBL for their generous support in developing this
curriculum.
Objectives - student should be able to:
1. Describe symbiotic relationships among organisms.
2. Describe the effects of a Wolbachia bacterial infection on insects.
3. Understand how to determine if an insect is infected with the Wolbachia bacteria based on DNA analysis.
4. Determine the closest match to a given DNA sequence using bioinformatics.
Introduction
Symbiosis
Organisms throughout the five kingdoms interact to varying degrees. These interactions or associations are called
symbioses (singular = symbiosis). Based on the nature of the interactions between two organisms, symbiotic
relationships can be characterized as one of three types: mutualism, commensalism, and parasitism.
If both organisms benefit from the relationship, it is considered mutualism. One example of this type of symbiosis
involves Lybia tesselata, or pompom crabs. These crabs house stinging sea anemones within their claws. When
approached by a threatening animal, the crabs wave around their claws (with stinging sea anemone “pompoms”),
which effectively deter the would-be predator. The crab gets protection, while the sea anemones get first dibs on
any food particles left over from the crab’s meals.
Commensalism describes a relationship in which one of the participating organisms benefits, while the other is
neither harmed nor helped. One example of this is plant epiphytes, such as orchids, that reside on the branches of
other trees. The orchids receive a high perch, which provides better access to sunlight and rainwater, while the
tree is neither harmed nor benefited by the presence of the orchid.
When one organism benefits at the expense of the other, the symbiosis is categorized as parasitism. In some
instances, the host is injured or sickened by the interaction; in others, the organism can be fatally harmed. In this
lab, we will be exploring one specific example of parasitism, in which parasitic bacteria in the genus Wolbachia, live
within the reproductive organs of insects and some other arthropods. This parasitic symbiosis has been shown to
cause population feminization through a variety of means, including male killing.
Despite convenient labels, symbiotic relationships are probably better characterized as falling within a continuum
between mutualism and parasitism. Some relationships may change status during the lifecycles of the organisms
involved, or there may be unobservable consequences of the relationship that cause miscategorization.
Additionally, a particular organism can have a parasitic relationship with one host and a mutualistic relationship with
another. This happens to be the case with Wolbachia, which is characterized as a parasite in arthropods; it is a
mutualist when inside filarial nematode hosts. Not only is Wolbachia a parasite, it is an endosymbiotic parasite,
meaning that it lives inside insect and nematode cells. For this laboratory, we will be focusing on Wolbachia the
parasite, which reside within the reproductive cells of arthropod species.
Wolbachia Infections
Unlike many parasitic organisms, which are transmitted to other hosts horizontally through contact with another
organism or the environment, Wolbachia can be transmitted both horizontally (though less commonly) and vertically
from parent to offspring, through the female’s eggs. Males do not pass the bacteria to their offspring.
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Wolbachia PCR
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Fall 2012
As an obligate intracellular bacterium, it must live inside another organism; it cannot reproduce outside of a host,
though it has been shown to survive short periods of time outside of a host cell. Therefore, to improve its rate of
transmission and success, Wolbachia has evolved several mechanisms by which it alters the sex of its host to
increase the number of females in the population. By reducing or eliminating the number of males in the
population, it prevents “dead end” transmission, ensuring that it is transferred with high efficiency to the next insect
generation.
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In some populations of insects, such as mosquitoes, beetles, wasps, moths, and fruit flies infected females can
only successfully reproduce with males that are infected with the same strain of Wolbachia bacteria. This type of
strategy is called “cytoplasmic incompatibility.” During spermatogenesis, factors from the bacteria modify the
sperm. This modification is “rescued” if the egg is also infected with the same strain of Wolbachia. The cause of
the incompatibility, at least in some insects, is due to DNA from the male and female being asynchronous or
uncoordinated during the initial stages of mitosis.
In other species, infection with the bacteria can cause females to reproduce parthenogenetically; eggs develop in
the absence of fertilization by the males. The resulting offspring are genetically identical to the mother; not only are
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they all female, they also all carry the Wolbachia infection . At the cytological level, the unfertilized (but infected)
eggs go through the first steps of mitosis, but the process stops at anaphase (the point at which the two sets of
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chromosomes separate), leading to a diploid nucleus . In the species in which Wolbachia infection induces
parthenogenesis, the unfertilized eggs would ordinarily develop into haploid males (with fertilized, diploid cells
developing into females). By interrupting the stage of mitosis of unfertilized, haploid cells at such a point that the
duplicated chromosomes fail to separate, a diploid female results.
In still other species, infection with Wolbachia causes fertilized eggs to develop as female, regardless of the
genotype. By suppressing the production of masculinizing hormones, genetically male embryos develop as
females. In the case of isopods (commonly referred to as “rolly pollies” or “sowbugs”), Wolbachia reside within a
male hormone-producing gland, reducing its effectiveness and thereby causing genetic males to develop as
phenotypic females. In other species, the mechanism of feminization is different and less well characterized.
Finally, in yet other insect orders, infection with Wolbachia simply causes male embryos to abort in early
development, allowing only genetically female embryos to hatch. In this way, the females have better access to
resources, which increases their survival rate.
Various studies have shown that cytoplasmic incompatibility and parthenogenesis can be “cured” with use of
antibiotics. Once fed antibiotics to eliminate the Wolbachia bacteria from their cells, the insects revert to the
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“normal” reproductive strategies . This finding was one of the main pieces of evidence supporting the hypothesis
that Wolbachia is the cause of these feminizing phenotypes. Please see the Appendix for a chart illustrating each
of the Wolbachia-induced phenotypes.
Evolutionary Consequences
Feminization of insect populations has a clear and positive effect on the Wolbachia population; the bacteria are
propagated efficiently. However, the effect of this sex-ratio distortion is less clear and almost certainly less positive
for the insect population.
In a situation in which females greatly outnumber males, it can be predicted that some females will never get the
opportunity to mate, thereby reducing the total number of offspring in a population. Furthermore, in species where
males usually compete for females and the females are choosy, Wolbachia-induced reduction of males causes a
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behavioral role reversal and mating favors the male that can most effectively attract the female . Through genetic
1
Meet the Herod bug (2001) Nature 412:12.
Stouthamer et al (1993) Molecular identification of microorganisms associated with parthenogenesis. Nature 361:66.
2
Stouthamer R, Kazmer DJ (1994) Cytogenetics of microbe-associated parthenogenesis and its consequences for gene flow in
Trichogramma wasps. Heredity 73:317.
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. Stouthamer R, Luck RF, Hamilton WD (1990) Antibiotics cause parthenogenetic Trichogramma to revert to sex. PNAS
87:2424
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Charlat S, Hurst GDD, Merçot H (2003) Evolutionary consequences of Wolbachia infections. Trends in Genetics 19:217.
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mutations, this alteration of typical behavior can be selected for thereby causing genetic variation in subsets of the
population.
In populations where the strategy has shifted from sexual reproduction to parthenogenesis, several consequences
can be expected. First, as populations become increasingly less diverse, they can be more easily decimated by
disease or pathogens. Secondly, prolonged reproductive isolation from other, non-infected groups of the same
type of insect can actually cause speciation by accumulation of deleterious mutations required for sexual mating.
Wolbachia-induced parthenogenesis has been implicated in speciation of one particular insect in which insects
“cured” of Wolbachia were unable and uninterested in mating.
Finally, evidence has shown that lateral transfer of DNA from Wolbachia to the host genome is widespread. This
has several consequences. First, the host can potentially begin producing proteins for the bacteria if the
transferred genes are transcribed by the host nucleus. Second, the random insertion of Wolbachia genes into the
host genome could have deleterious effects if inserted into a gene and possibly contribute to speciation. Finally,
some research suggests that Wolbachia may become another organelle, similar to mitochondria and chloroplasts.
Wolbachia Hosts: Classification
Worldwide, insect and other invertebrate numbers truly dwarf the human population. For every person on earth, it
is estimated that there are 3 billion insects! Even though we often think of animals as elephants, ducks, and
puppies, about 85% of all characterized animal species are insects, and since insects are often so small, this
number is probably underestimated, as many remain undiscovered.
Under the Linnaean classification system, living organisms are grouped according to similar characteristics. The
most broad grouping is called the “Kingdom,” and there are five categories: Plantae, Animalia, Monera, Protista,
and Fungi. Insects, as well as other invertebrates and vertebrates, fall in the category of Animalia. Within
Animalia, there are 21 phyla. Phylum Arthropoda encompasses invertebrates that have three body segments
(head, thorax and abdomen), a hard exoskeleton that is molted at intervals, and at least three pairs of jointed legs.
Within Arthropoda, there are seven major classes, including Insecta, Crustacea, Arachnida, and Diplopoda
(millipedes).
We often use the words ‘bugs’ and ‘insects’ to mean anything that is small and has more than four legs. However,
to be a true member of the class Insecta, the animal must have 6 legs (three pairs) and three body segments:
head, thorax and abdomen (see Figure 1.). Other scuttling creatures, such as rolly pollies (sowbugs) and spiders,
are not considered proper insects but land crustaceans and arachnids, respectively.
Head
Thorax
Abdomen
Antennae
Compound eye
Figure 1. Insect anatomy.
Legs
Within the Class Insecta, there are about 30 different orders of insects although there is some debate in the
scientific community regarding the exact number. Insects such as ants and bees fall in the Order Hymenoptera,
while butterflies and moths are part of the Order Lepidoptera. Each order has specific characteristics, and insects
can be classified using a dichotomous key. Additionally, there are many online resources for identifying and
learning more about insects. We recommend the following from Texas A&M, with helpful images of many species:
http://insects.tamu.edu/fieldguide/index.html.
Wolbachia has been identified in several classes within the phylum Arthropoda including insects, mites, spiders,
and crustaceans as well as some types of filarial nematodes (roundworms or threadworms, in the Phylum
Nematoda). Within the phylum Arthropoda, about 20% of all insect species are estimated to be
infected to varying degrees. Within some species, infection rates are as high as 90% or more.
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Wolbachia PCR
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In this lab, you will collect insects from your homes, schools, or other local areas. Using an online dichotomous
key, you will determine the order in which your insect belongs. We will only identify our insects to order; however,
every insect is classified into more specific groupings of genus and species.
Wolbachia Supergroups
Wolbachia species can be divided into eight supergroups (A-H), based on small variations in the 16S ribosomal
DNA and other genes. The A and B supergroups are most common in arthropods. Supergroups C and D are
typically found in filarial nematodes, while E is found in springtails (Order Collembola), F and H are found in
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termites (Order Isoptera), and G in some Australian spiders . Determining the supergroup to which a particular
Wolbachia species belongs is an active area of research as is identifying the variants of Wolbachia within each
supergroup.
Screening Insects for Wolbachia
Since Wolbachia is an obligate parasite, it is not possible to culture the bacteria from the insects to screen for their
presence. Therefore, we must use other methods to determine if a particular insect is infected. How can you
determine if a microscopic organism is living inside the cells of an insect? The answer is DNA! Every organism
has its own genome, or DNA sequence. Since organisms such as insects and bacteria have extremely distant
evolutionary ancestors, the DNA of each organism has had a lot of time to accumulate mutations or variations in
the DNA sequence. In fact, bacteria, such as Wolbachia, and insects are so different from each other that they are
classified in distinct Kingdoms (Monera and Animalia, respectively).
Scientists can take advantage of these differences at the DNA level by identifying regions of each organism’s DNA
that are unique to that type of organism. For this experiment, a gene was identified within the mitochondrial
genome that is almost identical in all insects. This gene is the cytochrome oxidase c gene (COI), the protein
product of which is involved in the electron transport chain and critical for the generation of ATP within mitochondria
organelles. Recall that the endosymbiotic theory states that mitochondria in eukaryotes are the result of an
ancient endosymbiotic relationship that persisted between bacteria (like Wolbachia) and a host. During coevolution of the cell and the endosymbiont, genes were transferred between the host cell nucleus and the DNA
within the bacteria, which effectively linked each organism’s survival to the other. Although, bacteria also have a
cytochrome oxidase c gene (despite lacking mitochondria themselves), it differs significantly from animals at the
DNA level. By searching for DNA sequence of COI specific to insects, we can effectively determine if we start with
insect DNA in our sample. In order to identify the presence of the bacterial DNA, a section of the 16S ribosomal
DNA is used that is unique to Wolbachia. Recall that ribosomes are responsible for translating RNA into protein.
While insects also have ribosomes, they are different from those found in bacteria at the DNA and protein level.
Furthermore, the section of ribosomal DNA that will be examined is unique to Wolbachia bacteria; that exact
sequence would not be found in any other bacteria. Therefore, means have been developed to identify DNA from
any insect and specific means to determine if Wolbachia bacteria DNA is present as well.
Finding the Microbes Within
In this lab, you will be extracting DNA from the insect you have collected. Insects are host to non-pathogenic
bacteria, just as humans and other animals are. Therefore, when we extract DNA from the insects, we will also
collect DNA from the bacteria that reside within their tissue. If the insect has been infected with Wolbachia, DNA
specific to that particular bacterium will be present. We will use polymerase chain reaction (PCR) to amplify two
DNA regions; 1) the COI gene from insects to verify we had a successful DNA extraction, and 2) the 16S ribosomal
DNA specific to Wolbachia to determine if the insect was infected. If the insect sample is infected with Wolbachia,
the DNA will be sent away for sequencing to determine the supergroup to which that particular Wolbachia variant
belongs.
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Bordenstein S and Rosengaus RB (2005) Discovery of a novel Wolbachia supergroup in Isoptera. Current Microbiology
51:393
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Wolbachia PCR
Student Guide
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Laboratory Exercise: Insect Identification
Using the insects collected prior to class, students will use an online dichotomous key to determine the Order to
which their insect sample belongs and prepare the insects for DNA analysis.
Objectives - student should be able to:
1. Understand the purpose of a voucher and create one for each insect collected.
2. Identify gross morphological characteristics of insects.
3. Use a dichotomous key to identify the Order in which your sample belongs.
Insect Identification and Vouchering
1. Place a Petri dish on the stage of your dissecting scope and
carefully squirt enough ethanol to cover the insects.
Note: Ethanol preserves the DNA for later lab activities.
Note: When not being used, insects should be kept in the freezer.

2. Using forceps, place one insect from your collection into the dish
with ethanol. Make sure your insect is submerged in the ethanol.

3. Examine the three main body regions of the insect: Head, Thorax,
and Abdomen.
Head Thorax
Abdomen

4. On the “Insect Identification Lab Mini-Report”, draw a detailed
picture of your insect, using colored pencils for accuracy and
record written observations of the head, thorax, and abdomen.
Name_______________________
Date_______________________
Insect Identification Lab Mini-Report
1.
Draw a detailed picture, using colored pencils if available, of
each of your morphospecies.

5. If a digital camera is available, or you have a digital microscope,
take several close-up images of the insect at various angles. This
image will serve as a voucher if you do not have a second,
identical insect.
Note: A voucher is a “copy” of your insect. It is used to further identify
the organism after the first one has been used DNA extraction.

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6. If you have a second, identical insect as a voucher, label a
microfuge tube with your name, date, and voucher. Be sure to
record your voucher number in your lab notebook. Fill the tube
halfway with ethanol and place your insect inside. Store this insect
in the freezer. You will NOT use the voucher insect for the DNA
extraction.

7. At the computer, open a browser window (Safari, Internet Explorer,
Firefox, etc.) and type the following URL into the address bar:
http://pick4.pick.uga.edu/mp/20q?guide=Insect_orders

8. After answering the questions in which you are confident, click on
any of the “Search” buttons to narrow the list of orders.
Definitions of insect features can be found in the left menu column
after clicking on the black and white pictures under each question.
Next, click on the “Simplify” button to eliminate unimportant
characteristics from your question list.
Click on image
for information
Proceed to answer more questions and use the “Search” and
“Simplify” buttons to determine the order of your insect.

9. Repeat these steps for each different insect your group has
collected.

10. Complete the “Insect Identification Lab Report” for your insect.
If you have collected the insect yourself, complete the chart
describing the habitat and life of each insect on the “Insect
Identification Lab Mini-Report”.
Name_______________________
Date_______________________
Insect Identification Lab Report
1.
Draw a detailed picture, using colored pencils if available, of
each of your morphospecies.

11. Place your insect in a 1.5 mL microfuge tube. Fill the tube halfway
with ethanol. Label the tube with the name, date, and voucher
number. Place the tube in a rack in the -20°C freezer until you are
ready to extract the DNA.

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Name________________________________________
Date _________________ Period_________________
Insect Identification Lab Report
1. Draw a detailed picture, using colored pencils if available, of your insect.
2. Complete the classification chart for your insect. For the “voucher label,” number your insect samples with your
two or three initials, the year, and a number (sequential, starting with 1). Example: JMJ2009-1. If you collected
more than one insect (including identical vouchers), continue the numbering (JMJ2009-2, JMJ2009-3, etc.). Be
sure to note if an insect is a voucher.
Scientific Name (Order)
Common Name
Voucher Label (s)
Notes
3. Research the habitat and life of your identified insect. Record notes in the following chart.
Insect
Common Name
Habitat
(trees, soil, etc.)
Diet
Life Cycle
Location Found
Interesting Facts
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Laboratory Exercise: DNA Extraction
In the following laboratory exercise, you will use several techniques to determine if the insect you have collected is
infected with Wolbachia bacteria. You will first isolate total DNA from the insect, which will also include Wolbachia
DNA, if present. The insect (or part of the insect) will be macerated in a buffer solution and the DNA will be
precipitated with isopropanol.
You will then use a process called polymerase chain reaction (PCR) to target and amplify two regions of DNA: an
insect region (present in all insects) and a Wolbachia region (if the insect is infected). Therefore, with each
successful DNA isolation, you should expect to observe the insect gene in all PCR reactions and the Wolbachia
gene in only the infected insects.. Finally, a process called gel electrophoresis will be used to visualize the
amplified products. The presence or absence of the amplified transgene element on your gels will provide
evidence as to whether the insect is infected with the Wolbachia parasite.
Objectives - student should be able to:
1. Successfully isolate DNA from insects.
2. Prepare a duplex PCR reaction to determine if your insect is infected with Wolbachia bacteria.
3. Cast an agarose gel to run your PCR products. After electrophoresis of your sample, you will then analyze
the photo results of the gel, comparing it with data from the rest of the class.
4. Explain your results, as well as why you used the positive and negative controls for this lab.
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Wolbachia PCR
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Im portant Laboratory Practices
a. Add reagents to the bottom of the reaction
tube, not to its side.
b. Add each additional reagent directly into
previously added reagent.
c. Do not pipet up and down, as this
introduces error. This should only be done
only when resuspending the cell pellet and not
to mix reagents.
d. Make sure contents are all settled into the
bottom of the tube and not on the side or cap of
tube. A quick spin may be needed to bring
contents down.
Keep reagents on ice.
a. Pipet slowly to prevent contaminating the
pipette barrel.
b. Change pipette tips between each delivery.
c. Change the tip even if it is the same reagent
being delivered between tubes. Change tip
every time the pipette is used!
Check the box next to each step as you complete it.
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Wolbachia PCR
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DNA Isolation from Insect Tissue
** Part 1: Preparation & Lysis of Insect Sam ple **
1. Obtain a clean 1.5 mL microfuge tube. Label the tube with the
insect voucher number.
Identification
on tube

2. Remove the insect from the alcohol and place it on a lab tissue
(Kimwipe) or paper towel to wick away any ethanol or isopropanol.

3. Obtain a ruler from the instructor. Measure the length of the
insect’s abdomen.
a. If the abdomen is less than 2 mm long and 2 mm wide, you
can use the entire abdomen.
b. If the abdomen is larger than 2 mm long and 2 mm wide, cut a
2 mm long section from the most posterior end of the abdomen.
c. If the entire insect is smaller than 2 mm, use the whole insect.

4. Place the insect or piece of insect into the labeled, clean microfuge
tube.
Note: Using an insect piece that is larger than 2mm will introduce
debris into the DNA and inhibit the PCR.
The piece of insect should sit about halfway to the 0.1 mL mark
A very small amount of the insect is all that’s needed! Remember,
you only need 1 strand of DNA.

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5. Add 200 µL of Lysis Buffer
Note: The Lysis Buffer breaks the cell and releases the DNA into
solution. DO NOT contaminate the stock Lysis Buffer solution!
200 µL Lysis Buffer

6. Macerate the insect by twisting the plastic micropestle for at least
1 minute or until only small bits of the chitinous exoskeleton
remain. You must use considerable downward and rotating force to
adequately macerate the insect tissue. If the insect gets stuck at
the bottom of the tube, use a clean pipette tip to dislodge it.
Note: You will not be able to completely macerate the chitinous
exoskeleton.

7. Once the insect is sufficiently macerated, add 800 µL Lysis Buffer.
Make sure the contents of the tube are thoroughly mixed by
vortexing or by “racking” the tube.
How much Lysis Buffer should you have in the tube at this point?
_________µL
800 µL
Lysis Buffer
Note: To “rack” your sample, be sure the cap of the tube is closed, hold
tube firmly at the top, and pull it across a tube rack 2-3 times.

8. Slide a cap lock onto your tube and place it in the 99°C heat block
or water bath for 5 minutes. Keep track of where your tube is in the
heat block or bath. The cap locks prevent tubes from popping open
due to vapor pressure.
Note: This step denatures proteins, including DNA-digesting enzymes.

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9. After heating, open tube briefly to release pressure, then close.
Vortex or shake tube to mix, and place in a balanced centrifuge.
Spin the tube for 5 minutes to pellet insect debris. Centrifuge
speed should be set to 10,000 x g (≈10,000 rpm).
Note: This step pellets insoluble material at the bottom of the tube
leaving DNA in the supernatant.
5 minutes at
10,000 x g

** Part 2: Rem oving Im purities **
During these steps, the DNA is located in the upper liquid portion, not the bottom pellet.
10. Obtain a second clean 1.5 mL microfuge tube. Label the tube with
your identification number.

11. Retrieve your tube from the centrifuge. There may be a noticeable
pellet at the bottom of the tube. Without disturbing this pellet,
withdraw 400 µL of the liquid (supernatant) from the centrifuged
tube and transfer it to the newly labeled tube. Discard the old
tube containing the pellet.
Note: You may notice an oily layer above the supernatant, which is
composed of residual lipids from the insect. If so, draw the liquid from
below the oily layer. Be careful not to disturb the pellet or any other
solid debris in the tube.
Transfer 400 µL of liquid
pellet at bottom

12. Add 40µl of 5M NaCl to the tube containing the 400µl of liquid that
you removed. Shake the tube a few times to mix. Incubate on ice
for 5-10 minutes. Solution may become cloudy.
40 µL of NaCl
5 minutes
on ICE
Note: The NaCl binds to detergents in the Lysis Buffer that we need to
remove in order to have a clear DNA sample.

13. Place tube with NaCl into centrifuge and spin again as described in
step 9 above.
5 minutes at 10,000 x g

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** Part 3: Isolating the DNA **
During these steps, the DNA is located in the pellet at the bottom of the tube.
14. Obtain a third clean 1.5 mL microfuge tube. Label the tube with
the word “DNA” and your identification number.

15. Retrieve your tube from the centrifuge. There may be a noticeable
pellet at the bottom of the tube. Repeat what you did in step 11, but
this time take 300 µL of the liquid from the top of the tube. Be
careful not to take anything from the pellet at the bottom of the tube.
Transfer 300 µL of liquid
to new tube

16. Add 400 µL isopropanol to your new tube of the transferred liquid.
Mix contents by inverting your tube several times, or vortex for a
few seconds.
400 µL of isopropanol
Note: The isopropanol precipitates DNA and makes it settle to the
bottom of the tube.
Pellet at bottom

17. Place the tube with the isopropanol mixture in a balanced
centrifuge. Centrifuge speed should be set to 10,000 x g
(≈10,000 rpm).
VERY IMPORTANT: Orient the hinge of the tube to point
outward and away from the middle of the centrifuge.
Spin at top speed for 5 minutes.
Note: Nucleic acids (DNA) will pellet at the bottom-side of the tube
under the hinge during centrifugation.
5 minutes

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18. Retrieve your tube from the centrifuge. Carefully pour the liquid out
of the tube. Tap the mouth of the tube lightly onto a clean paper
towel to remove the liquid on the lip of the tube. Spin quickly again
in the centrifuge to pool the rest of the liquid and use a pipette to
completely remove all of the supernatant. Aim the tip away from
the pellet to remove the liquid. DO NOT disturb the pellet.
Note: The DNA is now in the bottom of the tube. The DNA pellet may
leave a teardrop-shaped mark or may appear as minute speckles on
the hinge-side of the tube. Do not worry if there is no visible pellet.
DNA

!
19. Air-dry the pellet for about 5-10 minutes to evaporate any
remaining isopropanol. Keep the cap open.
Note: To speed up the evaporation process, place tubes on a heat
block set at about 50-70°C. Keep caps open and monitor for
evaporation. If most of the liquid has been removed from the tubes
beforehand, this should take less than 5 minutes.

20. IMPORTANT: Use filter tips for this step to avoid
contamination of the pipette barrel.
200 µL TE/RNase
Add 200 µL of TE/RNase buffer to your tube. Scrape the side of
the tube where the pellet is (or should be) with the tip to facilitate
resuspension. Pipette up and down several times to collect DNA
accumulated on the area underneath the hinge.
Note: TE buffer stabilizes the DNA. RNase destroys any isolated RNA
that might interfere with the PCR reaction.

21. Observe your DNA. If it is clear, proceed to the next step. If it is
cloudy or if there is a lot of debris, add another 200 µl TE/RNase.
22. After the resuspension, place your tube in a centrifuge. Balance
and spin the tube for 1 minute to pellet any particulates that did not
dissolve in solution.

Note: This is your isolated insect (and possibly Wolbachia) DNA! It
may contain nucleases that can degrade the DNA at room temperature.
Keep the DNA tubes on ICE to limit any leftover nuclease activity.
23. Place your DNA tube in the class rack. Your teacher will FREEZE
your isolated DNA until you are ready to prepare your PCR
amplification.
Note: Your plant DNA extract may contain nucleases that can degrade
the DNA at room temperature. Keep the DNA tubes on ICE to limit any
leftover nuclease activity.
!
1 minute


24. Be sure to clean micropestiles for re-use and soak in ethanol to
sterilize. This will reduce cross-contamination when they are used
by the next class.

14
Wolbachia PCR
Student Guide
Fall 2012
Laboratory Exercise: Polym erase Chain Reaction
Objectives - student should be able to:
1. Explain the importance of each component of PCR and compare it to in vivo DNA replication.
2. Associate the temperature changes with the cycling steps of PCR.
3. Understand the concept of a duplex PCR reaction.
4. Explain the purpose of the two different sets of primers used in this duplex PCR.
The polymerase chain reaction (PCR) is a technique used by scientists to rapidly multiply specific segments of
DNA in a small tube. Essentially, PCR reproduces some of the mechanisms of cellular DNA replication and thus
has the capacity to churn out millions of copies of a targeted DNA segment. This PCR amplification is used in
many aspects of science including forensics, human identification, genetic disease diagnostics, and in the cloning
of rare genes. One of the reasons PCR has become such a powerful technique is that it does not require large
amounts of DNA; a drop of blood at a crime scene, tiny quantities of food matter, a small leaf of a plant, or the back
of a licked postage stamp usually has sufficient DNA for PCR amplification.
There are some essential reaction components and conditions needed to amplify DNA by PCR. First, it is
necessary to have a sample of DNA containing the segment you wish to amplify. This DNA is called the template
because it provides the base sequence to be duplicated during the PCR process. Along with template DNA, PCR
requires two short single-stranded pieces of DNA called primers. These are usually about 20 bases in length and
are complementary to the opposite strand of the template. Each primer is designed to sit down at one end of the
target DNA segment being amplified. Primers attach (anneal) to their complementary sites on the template and are
used as initiation sites for synthesis of new DNA strands. Deoxynucleoside triphosphates (dNTPs) containing
the bases A, C, G, and T are also added to the reaction. The enzyme DNA polymerase binds to one end of each
annealed primer and strings the dNTPs together to form a new DNA chain complementary to the template. The
++
DNA polymerase enzyme requires the metal ion magnesium (Mg ) for its activity. It is supplied to the reaction in
the form of MgCl2 salt. A buffer is used to maintain an optimal active pH level for the DNA polymerase.
PCR is accomplished by cycling a reaction through several temperature steps. In the first step, the two strands of
the template DNA molecule are separated, or denatured, by exposure to a high temperature (usually 94° to 96°C).
Once in a single-stranded form, the bases of the template DNA are exposed and are free to interact with the
primers. In the second step of PCR, called annealing, the reaction is lowered to a temperature usually between
37˚ to 65˚C. At this lower temperature, stable hydrogen bonds can form between the complementary bases of the
primers and the template. Although human genomic DNA is billions of base pairs in length, the primers require only
seconds to locate and anneal to their complementary sites. In the third step of PCR, called extension, the reaction
temperature is raised to an intermediate level (65˚ to 72˚C). During this step, the DNA polymerase starts adding
nucleotides to the ends of the annealed primers. These three phases are repeated over and over again, doubling
the number of DNA molecules with each cycle. After 25 to 40 cycles, millions of copies of desired DNA are
produced. The PCR process taken through four cycles is illustrated on the following page (Figure 2).
For this reaction, there will be two sets of primers: one set that is specific for the cytochrome c oxidase gene (COI)
from the insect mitochondrial genome, and one set that is specific for the Wolbachia-specific bacteria ribosomal
DNA (Wspec). Amplification using the COI primers results in a band 709 bp long, while amplification with the
Wspec primers results in a band 438 bp long. The two independent PCR reactions will occur within the same tube,
one amplifying a region of bacterial DNA and the other amplifying a region of the insect DNA. Reactions that
include two sets of primers, for the purpose of amplifying two different regions, are called “duplex reactions”.
15
Wolbachia PCR
Student Guide
Fall 2012
Illustration of the Polym erase Chain Reaction
Figure 2. The first four cycles of the polymerase chain reaction.
First Cycle of PCR
Third Cycle of PCR
Second Cycle of PCR
Fourth Cycle of PCR
PCR images from Life Technologies, formerly Applied Biosystems, Inc.
An excellent animated tutorial showing the steps of PCR is available at the DNA Learning Center web
site. http://www.dnalc.org/ddnalc/resources/pcr.html. Note: You may need the Macromedia Flash plugin to view this animation.
16
Wolbachia PCR
Student Guide
Fall 2012
Polym erase Chain Reaction
1. Obtain a tiny PCR tube. Label it with your voucher number, just
under the lip.
Note: Keep your PCR tube on ice when setting up the reaction.

2. Pipet 20 µL of Master Mix into your PCR tube.
20 µL of
Master Mix

3. Change your pipet tip and add 20 µL of Primer Mix into your PCR
tube.
20 µL of
Primer Mix

4. With a new tube, add 10 µL of your extracted DNA into your PCR
tube. What is the total volume in your tube? ___ µL.
10 µL of
DNA
Note: Make sure that all the liquids are settled into the bottom of the
tube and not on the side of the tube or in the cap. If not, you can give
the tube a quick spin in the centrifuge. Do not pipette up and down, it
introduces error.

5. Setting up the controls:
a) Two students will be asked to set up the positive control
reactions for the class. They will use the positive control DNA
provided in the kit. There should be enough PCR sample for
one lane on each gel.
Control
Master
Mix
Primer
mix
DNA
+
20 µL
20 µL
10 µL +C DNA
-
20 µL
20 µL
10 µL sterile H20
b) Another two students will set up negative control reactions for
the whole class. They will use sterile water. There should be
enough PCR sample for one lane on each gel.

6. Check the volume of your PCR tube by comparing it to a reference
PCR tube with 50 µL in it. It should be near the thermal cycler, set
by your teacher.
50
50#μL#
Note: If the volume of your tube does not match, see your instructor to
troubleshoot. You may need to set up a new reaction.
Reference Tube 
PCR Tube
7. Place your reaction into the thermal cycler and record the location
of your tube on the grid provided by your teacher.
1
A
B
C
2
3
4
1123
5
6
7
828
1027
6777
9305

17
Wolbachia PCR
Student Guide
Fall 2012
8. The cycling protocol for amplification of Wolbachia PCR:
Thermal cycler Instrument displaying
program parameters
1) 95°C hold for 2 minutes
2) 30 cycles of:
94°C for 30 seconds
55°C for 45 seconds
72°C for 1 minute
3) 72°C hold for 10 minutes
4) 4°C hold, ∞ infinity
18
Wolbachia PCR
Student Guide
Fall 2012
Agarose Gel Electrophoresis
To determine whether or not your insect contains Wolbachia, you will need to visualize the products of your
amplification. This will be done using a process called gel electrophoresis in which electric current forces the
migration of DNA fragments through a special gel material. Since DNA is negatively charged, it will migrate in an
electric field towards the positive electrode (Figure 2). When electrophoresed through a gel, shorter fragments of
DNA move at a faster rate than longer ones.
Figure 2. Side view of an
agarose gel showing DNA
loaded into a well and the
direction of DNA fragment
migration during
electrophoresis.
The gel material to be used for this experiment is called agarose, a gelatinous substance derived from a
polysaccharide in red algae. When agarose granules are placed in a buffer solution and heated to boiling
temperatures, they dissolve and the solution becomes clear. A comb is placed in the casting tray to provide a mold
for the gel. The agarose is allowed to cool slightly and is then poured into the casting tray. Within about 15 minutes,
the agarose solidifies into an opaque gel having the look and feel of coconut Jell-O™. The gel, in its casting tray, is
placed in a buffer chamber connected to a power supply and running buffer is poured into the chamber until the gel
is completely submerged. The comb can then be withdrawn to form the wells into which your PCR sample will be
loaded.
Loading dye is a colored, viscous liquid containing dyes (making it easy to see) and sucrose, Ficoll, or glycerol
(making it dense). To a small volume of your total PCR reaction, you will add loading dye, mix and then pipet an
aliquot of the mixture into one of the wells of your agarose gel. When all wells have been loaded with sample, you
will switch on the power supply. The samples should be allowed to electrophorese until the dye front (either yellow
or blue, depending on the dye used) is 1 to 2 cm from the bottom of the gel. The gel can then be moved, stained
and photographed.
Calculations for Preparing 2% Agarose Gel
You will need a 2%, mass/volume agarose gel for electrophoresis of your PCR products. If your agarose gel
casting trays holds 50 mL, then how much agarose and buffer would you need? The definition of m/v % in biology
is grams (mass) / 100 mL (volume). Therefore, for 2% agarose, it will be 2 g /100 mL buffer.
Step 1: Calculate the mass of agarose needed for 50 mL total volume of agarose solution.
2g
Xg
=
100 ml
X = 1 gram
50 ml
Step 2: Calculate the amount of buffer needed to bring the agarose solution to 50 mL. By standard definition, 1
gram of H2O = 1 mL of H2O. The amount of buffer for the 2% agarose solution will be 49 mL (50 mL – 1 mL (1 gram
of agarose)).
Note: If using an Invitrogen E-Gel (pre-cast agarose gel) and E-Gel PowerBase, please follow
supplementary instructions located on page 22.
19
Wolbachia PCR
Student Guide
Fall 2012
Electrophoresis of Am plified DNA
1. Retrieve your PCR tube and place it in a balanced configuration in
a microcentrifuge. Spin it briefly (10 seconds) to bring the liquid to the
bottom of the reaction tube.
Note: Make sure the centrifuge adapters are in place before
putting the tiny PCR tube into the centrifuge rotor.
2. If you are NOT performing DNA sequencing:
Add 5 µL of loading dye to your PCR tube.
If you plan to sequence your DNA:
Remove 20 µL of your PCR sample and dispense into a new
tube. Add 2 µL of loading dye to it.
Note: your PCR sample can’t contain loading dye for sequencing.
3. Carefully load 15 to 20 µL of the DNA/loading dye mixture into a
well in your gel. Make sure you keep track of what sample is being
loaded into each well.
Note: Avoid poking the pipette tip through the bottom of the gel
or spilling sample over the sides of the well. Use a new tip for
each sample.
4. One student (or the instructor) should load 5-10 µL of 100 bp ladder
(molecular weight marker) into one of the wells of each gel.
5. When all samples are loaded, attach the electrodes from the gel box
to the power supply. Have your teacher check your connections and
then electrophorese your samples at 150 Volts for 25–40 minutes.
6. After electrophoresis, the gels will be ready to stain and photograph.
20
Wolbachia PCR
Student Guide
Fall 2012
Staining and Photographing Agarose Gels
The PCR products separated on your agarose gel are invisible to the naked eye. If you look at your gel in normal
room light, you will not be able to see the amplified products of your reaction. In order to “see” them, we must stain
the gel with a fluorescent dye called ethidium bromide (EtBr). Molecules of ethidium bromide are flat and can
intercalate, or insert, between adjacent base pairs of double stranded DNA (Figure 3). When this interaction
occurs, they take on a more ordered and regular configuration causing them to fluoresce under ultraviolet light
(UV). Exposing the gel to UV light after staining, allows you to see bright, pinkish-orange bands where there is DNA
(figure 4).
Figure 3. Ethidium bromide
molecules intercalated
between DNA base pairs.
Your teacher may stain your agarose gel and take a photograph for you so that you may analyze your Wolbachia
results.
Gel staining is done as follows:
1. Place the agarose gel in a staining tray.
2. Pour enough ethidium bromide (0.5µg/ mL) to cover the gel.
3. Wait 20 minutes.
4. Pour the ethidium bromide solution back into its storage bottle.
5. Pour enough water into the staining tray to cover the gel and wait 5 minutes.
6. Pour the water out of the staining tray into a hazardous waste container and place the stained gel on a
UV light box.
7. Place the camera over the gel and take a photograph.
8. Check with your district on how to dispose of hazardous waste liquid and solids.
CAUTION: Ethidium bromide is considered a carcinogen and neurotoxin. Always wear gloves and
appropriate PPE (personal protective equipment) like safety glasses when handling. Students should
NEVER handle EtBr.
CAUTION: Ultraviolet light can damage your eyes and skin. Always wear protective clothing and UV safety
glasses when using a UV light box.
Figure 4. After staining an agarose gel
with ethidium bromide, DNA bands are
visible upon exposure to UV light.
21
Wolbachia PCR
Student Guide
Fall 2012
Instructions for Using Invitrogen E -Gel and E-Gel PowerBase
Before getting started, determine which kind of power base you have. There are two options:
Option 1: A red E-Gel base, which is connected to
a single power cable with an adaptor. It can be
plugged directly into an electrical outlet.
Option 2: A black E-Gel base, which has two power
cables: a red one that leads to the anode (+), and a black
one that leads to the cathode (-). It needs to be plugged
into a power supply.
Directions for Option 1: Red E -Gel base
1. Remove E-Gel from the package and insert it with the
comb in place into the base, right edge first. The Invitrogen
logo should be located at the bottom of the base. Press firmly
at the top and bottom to seat the E-Gel into the base.
Plug the base into an electrical outlet using the adaptor plug on
the base. A steady, red light illuminates if the gel is correctly
inserted. Wash hands.
Note:
Wear gloves when assembling the E-Gel, which
contains ethidium bromide. Small amounts of buffer may
emerge from the wells during assembly.
2. Remove combs from the E-Gel cassette and dispose of as
EtBr waste.
Load 20µl of DNA ladder.
Load 20µl of prepared sample into each well.
Load 20µl of water into any remaining empty wells.
Press the 30-minute button to start the run.
Light will turn green while run is in progress.
E-Gel automatically stops but will continue to beep
until unit is turned off.
Note: Leaving empty wells will cause the E-Gel to run
unevenly. Remember to add water to empty wells.
22
Wolbachia PCR
Student Guide
Fall 2012
3. The run can be interrupted at any time. 20-30 minutes will
be sufficient. Loading dye should move at least halfway
through the length of the gel, as shown.
Run progress can be checked on a transilluminator. If band
separation is not complete, simply return E-Gel to base and run
longer. Alternatively, entire unit can be placed under the
transilluminator to check progress.
The E-Gel already contains ethidium bromide. Gel staining is
not necessary. At the end of the run, proceed directly to
imaging the E-Gel.
Directions for Option 2: Black E-Gel base
1. Remove E-Gel from the package and insert it with the
comb in place into the base right edge first. The Invitrogen
logo should be located at the bottom of the base. Press firmly
at the top and bottom to seat the E-Gel into the base.
Plug the two cables located on the black base into an
electrophoresis power supply. The red (+) cable connects to
the red slot and the black (-) cable connects into the black slot.
Wash hands.
Note:
Wear gloves when assembling the E-Gel, which
contains ethidium bromide. Small amounts of buffer may
emerge from the wells during assembly.
3. Remove combs from the E-Gel cassette. Make sure power
supply is turned off.
Load 20µl of DNA ladder.
Load 20µl of prepared sample into each well.
Load 20µl of water into any remaining empty wells.
Turn on power supply and run E-Gel at 60-70 volts.
Note: Leaving empty wells will cause the E-Gel to run
unevenly.
!
4. The run can be interrupted at any time. 20-30 minutes will
be sufficient. Loading dye should move
at least halfway through the length of the E-Gel.
Run progress can be checked on a transilluminator.
If band separation is not complete, simply return E-Gel to base
and run longer. Alternatively, entire unit can be placed under
the transilluminator to check progress.
The E-Gel already contains ethidium bromide. Gel staining is
not necessary. At the end of the run, proceed directly to
imaging the E-Gel.
23
Wolbachia PCR
Student Guide
Fall 2012
Results
By examining the photograph of your agarose gel, you will determine whether or not the insect you identified is
infected with the Wolbachia bacteria. The primer mix you used when setting up your PCR contained two sets of
primers. One set amplified the COI gene in insects. If the DNA from your insect was isolated successfully, PCR
amplification of this gene from the insect genome will produce a fragment 709 bp in length. The second set of
primers amplifies a Wolbachia-specific region of bacterial ribosomal DNA 438 bp in length. If the insect is infected
with Wolbachia bacteria, PCR amplification of the ribosomal DNA will result in a second band in the same lane on
the gel.
709 bp
438 bp
Figure 6. Agarose gel of insects with and without Wolbachia bacterial infections. A 100 base pair ladder is loaded
in the first lane as a size marker in which the smallest band is 100 bp and every subsequent band is 100 bp larger.
The 500 bp band and the 1,000 bp band are purposely spiked for greater fluorescent intensity than are the other
bands of the ladder when stained with ethidium bromide. The positive control lane shows the two expected bands.
Lanes #178, #180, and #183 show definite, strong positive bands for Wolbachia. Lanes #177 and #179 are also
positive, but the bands are fainter most likely due to a lower titer of bacteria
24
Wolbachia PCR
Student Guide
Fall 2012
Uploading Class Results to the BABEC Wolbachia M ap
The new BABEC Wolbachia Map has been released! Please join us in this collective effort to document Wolbachia
prevalence in the Bay Area. As more data is collected, future classes will be able to see where the Wolbachia “hot
spots” are in the area. We also hope to share this data with researchers in the field to aid them in their
investigations. It’s a great opportunity for your students to participate in real world science and collaborate like a
real scientist.
Please upload your class data to the map! Please note that only teachers can create accounts and upload data,
but students and other users can observe and export data from the map.
Instructions for Uploading Wolbachia Data to the Map
1. Click on the map link or copy the address into your web browser: http://wolbachiamap.herokuapp.com/
2. Click on “Log In” in the top right corner of the page. If you don’t already have an account, click on “Create New
Account” and fill in the required fields. New accounts are for BABEC teachers only, and require administrator
approval before you can log in.
3. Once your account has been activated, you can log in as an approved user. Teachers can use the same
account for all classes and only one account is needed per teacher. Each class dataset will require a date and
class name for tracking.
4. Click on the “Add/Edit Data” for instructions on how to upload your class data. It will be helpful to have
completed the data table (located on page XV of the teacher guide) before uploading.
5. Click on “Explore Data” and “Explore Map” to look at the full dataset for the Bay Area. These tools allow you to
select specific regions or types of insects to look at. Feel free to export data, make graphs, and investigate the
map.
6. Although only teachers can upload data, the map is publicly available for browsing. The map is interactive and
can be viewed by students, teachers, and the general public.
This map was developed in collaboration with computer science students from UC Berkeley, and is a
project of The Berkeley Group. We thank them for their generous support
25
Wolbachia PCR
Student Guide
Fall 2012
How to subm it PCR sam ples for sequencing through CSU East Bay

1. Obtain a clear photo of your class gel results. This photo is
required for the processing of your samples at CSUEB. Label the
lanes in sequential order, using only numbers, beginning with 1.
2. Spin down the original PCR tube. With a fresh tip, remove 10
µL of the PCR product and place in a new tube.
3. Label the new tube with the corresponding number from the gel
photo.
1
!
!


4. The contact person at CSU East Bay is Dr. Chris Baysdorfer.
Contact him via email at the time of shipment to confirm. Use the
following email address:
chris.baysdorfer@csueastbay.edu
Include the following class information:
- your name
- school
- number of samples
- class name (AP Bio, etc.).
5. Package your samples and labeled gel photo in a zip-top
plastic bag. Place in a styrofoam box with a few cooler packs.
26
Wolbachia PCR
Student Guide
Fall 2012
6. Ship OVERNIGHT delivery to:
Professor Chris Baysdorfer
Department of Biological Sciences
California State University, East Bay
Hayward, CA 94542
Phone: (510) 885-3459
IMPORTANT: Do not ship samples on Fridays.
7. The sequences will be delivered to you via email. This should
take 5-7 business days, but confirm with Dr. Baysdorfer at time of
shipment. Samples are run free of charge!
8. You will receive the sequences from CSU in the form of a
“Trace File”. They will have the same sample names that you
submitted, but will end in “.ab1”. The .ab1 files need a special
software program to be opened. Programs can be downloaded for
free:
“4Peaks” for MAC
For MAC
http://www.mekentosj.com/science/4peaks
For PC:
https://products.appliedbiosystems.com/ab/en/US/adirect/ab?cmd
=catNavigate2&catID=600583&tab=Overview
9. Using these programs, you can visualize and edit the color
chromatogram of each sequence. You can also export the
sequences as text files, which will be needed for further analysis
using the Sequence Server at the CSHL DNA Learning Center.
See instructions below, “Mitochondrial sequence comparisons”,
starting on page 32.
Note: Feel free to contact the Manager of Education Programs at
BABEC if you need help using these programs or exporting the
text files. Contact information can be found at www.babec.org
10. Sequences can be shared with scientists at the Wolbachia
Project at Wood’s Hole Marine Biological Laboratory:
http://discover.mbl.edu.
Contact the Manager of Education
Programs at BABEC for details on this exciting option.
27
“Peak Scanner” for PC
Sequence chromatogram:
Sequence text file:
Share your data with scientists at
The Wolbachia Project!
Wolbachia PCR
Student Guide
Fall 2012
Bioinform atics Exercise: Sequence Taxonom y
Objectives:
To become more familiar with the National Center for Biotechnology Information (NCBI) website resources.
To provide an introduction to the number and diversity of nucleotide sequences in the NCBI database.
The National Center for Biotechnology Information (NCBI) website (http://www.ncbi.nlm.nih.gov/) is an incredible
resource for molecular biology. According to their mission:
“NCBI has been charged with creating automated systems for storing and analyzing knowledge
about molecular biology, biochemistry, and genetics; facilitating the use of such databases and
software by the research and medical community; coordinating efforts to gather biotechnology
information both nationally and internationally; and performing research into advanced methods of
computer-based information processing for analyzing the structure and function of biologically
important molecules.” --NBCI website
There is a veritable treasure trove of information on the NCBI website; from searchable databases of published
literature (PubMed) and nucleotide sequence information (GenBank), to powerful tools to analyze those sequences
(BLAST, Entrez Gene, ORF Finder, etc.) and mine the vast amounts of compiled data (Entrez, BLAST, Taxonomy
Browser, etc.). While we can only touch the tip of the iceberg here, it is worth the time to browse around on your
own. Here, we will be exploring Taxonomy Statistics to give a feel for the breadth of information available on NCBI.
Sequence Taxonom y
1. Open up a browser and type in the following address into
the URL bar:
www.ncbi.nlm.nih.gov/
Select ‘Taxnonmy’ on the LEFT menu bar

2. In the CENTRAL menu, about halfway down, select
‘Taxonomy Statistics’, which is located under the ‘Tools’
heading.

28
Wolbachia PCR
Student Guide
Fall 2012
3. In the table that opens up, select the year 2011 (in the
‘dates’ section below the table). Find the number of
Bacterial Species currently represented in the database.
______________________
Now, click on the year 2002. How many Bacterial Species
were added in that year?
_______________________
What a difference a few years makes!

4. Interestingly, sequence data from extinct organisms is even
listed in the GenBank database. Let’s look for a gene
sequence from a 120 million year old insect preserved in
amber!
From the same page as above, select ‘Extinct Organisms’
from the LEFT menu bar’. Scroll all the way down the page
to find ‘Insects’ and select ‘Libanorhinus succinus’

6. What are some other organisms that belong to this phylum of animals? _______________________________
___________________________________________________________________________________________
7. Can you think of any body traits that these organisms have in common? ______________________________
___________________________________________________________________________________________
8. How many ‘nucleotide’ sequences have been deposited into
the Entrez records for this organism (top right corner of
page)?
__________________________
Click on the number to bring up the record. What is the
name of the gene that was sequenced from this organism?
__________________________
How many nucleotide base pairs does this entry have?
29

Wolbachia PCR
Student Guide
Fall 2012
9. Click on the link for the complete record of this sequence. A
lot of information is listed, to provide scientists with as much
data about the sequence as possible.
At the bottom of the screen, you will find the nucleotide
sequence itself.

10. Click on the PUBMED number above the sequence. This
link will take you directly to the published journal article that
describes the discovery of this nucleotide sequence.
Select the ‘NCBI’ link in the top left corner of the screen to
return to the main home page.

30
Wolbachia PCR
Student Guide
Fall 2012
Bioinform atics Exercise: Sequence Searching Using BLAST
Objectives:
To obtain sequence information from the NCBI database that matches your Wolbachia sequence using the BLAST
(Basic Local Alignment Search Tool) at NCBI.
To determine strain information about the Wolbachia bacteria identified in your insect sample.
Introduction
When an organism’s genes (or proteins, RNA, or whole genome) is sequenced, the information is often uploaded
by scientists to GenBank, a repository for genetic sequences on the public National Center for Biotechnology
Information (NCBI) database (http://www.ncbi.nlm.nih.gov/). Each sequence record is annotated with the
organism’s name, information about the researcher, publications about the sequence, and other useful information
(For
a
detailed
description
of
the
components
of
the
sequence
records,
see
http://www.ncbi.nlm.nih.gov/Sitemap/samplerecord.html). In addition to GenBank, the NCBI website contains an
incredible amount of information, but for the purposes of this lab, we are only going to focus on one tool: BLAST.
If you have sequence information in hand, or are just interested in looking at sequence data other groups have
published, one of the most useful tools is the BLAST search. BLAST stands for Basic Local Alignment Search
Tool, and is used to identify regions of local similarity between two different nucleotide sequences. Since all
organisms have the same genetic code of A, T, C and G’s, the sequence and arrangement of these nucleotides
can be compared to look for similarities and differences between organisms. BLAST is also frequently used for
determining the identity of an unknown DNA sequence. If you have protein sequence instead of nucleotides, you
can also search and compare amino acid (protein) sequences. In either case, the sequence that you input into the
search is called the “query” and the resulting sequences are called the “subject”. If your class sent away DNA to be
sequenced and you have received back a digital file containing the sequence, then your “query” will be that
sequence. If you did not, you can use the one provided.
In this module, you will do some short exercises to get a feel for how to use BLAST. Once you know how to use
the tool, you can try performing a search on the sequence information about your own sample. If you don’t have
your own data, go to (www.babec.org/node/93) to obtain a sequence with which to search.
Sequence Searching Using BLAST
1. Open up a browser and type in the following address into
the URL bar:
www.ncbi.nlm.nih.gov/
Select ‘BLAST’ in the RIGHT menu bar.

31
Wolbachia PCR
Student Guide
Fall 2012
2. Under the ‘Basic BLAST’ heading, click on the link that says
‘nucleotide BLAST’.
The nucleotide BLAST is specifically for comparing
nucleotide sequences to other nucleotide sequences. This
will pull up a page that looks like the image to the right.
Notice the other options under Basic BLAST.

3. In the ‘Query Sequence’ box, input a random sequence of
A, T, C, and Gs, so that you fill the top line. You have just
entered a ‘query’.

4. Below the ‘Query Sequence’ box, under the heading
‘Choose Search Set’, click the button for ‘Others (nr etc.)’.
This is very important to ensure we are searching within the
whole database, and not a subset of sequences.
Note: “nr” stands for “non-repetitive”.
Now, click on the big ‘BLAST!’ button at the bottom of the
page. A new window will appear.
Note: It may take several seconds for the BLAST to complete.

5. Did your sequence produce a significant match or hit?
If yes, how many records matched your random sequence?
Note: a significant match is one in which the E value is less than E-10.
6. Click on the ‘Other reports > Search Summary’ link near
the top of the page. This will pull up information about your
specific search.
How many sequences did it search in the database?
How many nucleotides (letters) did it search in the
database?
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
Wolbachia PCR
Student Guide
Fall 2012
7. At the top of the page, click on ‘HOME’ to return to the
BLAST homepage. As in Step #3, click on ‘nucleotide
BLAST’ under the ‘Basic BLAST’ heading.

8.
In the text box, enter the sequence of nucleotides listed in
the Appendix, or go to www.babec.org/node/93 and click on
the link to download the sequence file.
Copy and paste the sequence into the text box.
Be sure the ‘Other (nr etc.)’ button is selected under
‘Choose Search Set’. Hit the ‘BLAST!’ button.

9. You should pull up a results page that looks similar to the
image on the right. Each line indicates a sequence within
the database that has similarity to your query. Complete
the questions, below, before continuing.

10. How many nucleotides was your query? ________________________________________________________
11. What is the accession number for the first hit?____________________________________________________
12. What is the ‘E value’ for the first hit?___________________________________________________________
13. What is the most likely identity of this sequence? (Click on the blue ‘Accession’ link to the left of the top hit.)
___________________________________________________________________________________________
14. Is there any information about the Wolbachia strain? If so, what is it?_________________________________
15. What is the title of the scientific publication that reported this sequence?______________________________
__________________________________________________________________________________
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Wolbachia PCR
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16. Select ‘Home’ at the top left of the BLAST page.
Now select ‘nucleotide BLAST’ under the ‘Basic BLAST’
heading.

17. Enter in the first 20-25 base pairs of the Wolbachia
sequence you entered in step 10.
As before, click the radio button for ‘Other (nr etc)’ and then
hit ‘BLAST!’

18. You should pull up a results page that looks similar to the
image on the right. Each line indicates a sequence within
the database that has similarity to your query

19. What is the ‘E-value’ for the first hit? __________________________________________________________
20. Is this ‘E-value’ more or less significant than the longer sequence you BLAST’ed in step 10? (Remember,
numbers raised to a large negative power are smaller than numbers raised to a small negative power.)
___________________________________________________________________________________________
21. From the two BLAST searches, what can you deduce about how the length of the query sequence affects your
confidence in the sequence search?
__________________________________________________________________________________
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Wolbachia PCR
Student Guide
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Wolbachia PCR
Student Guide
Fall 2012